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10.4: Exercise 1 - Plasmid isolation with the ZyppyTM kit - Biology

10.4: Exercise 1 - Plasmid isolation with the ZyppyTM kit - Biology


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Obtain the plasmid-bearing bacterial cells

1. Collect the three bacterial cultures that your group has been assigned. The bacteria have been transformed with plasmids containing either an S. cerevisiae MET gene, its S. pombe ortholog or bacterial LacZ. Each culture contains 0.6 mL Luria Bertani (LB) medium with 100 μg/mL ampicillin. Cultures were grown overnight at 37 degrees C, and the cell density is expected to be 3-4 X 109 cells/mL.

What is the purpose of the ampicillin? How does it work?

Alkaline lysis of bacterial cells harboring the plasmids

2. Add 100 μL of 7X Blue Zyppy Lysis buffer to the tube. Mix the buffer and cells by gently inverting the tube 4-6 times. Be gentle! Too much mechanical stress could fragment the bacterial chromosomal DNA and contaminate your plasmid preparation. The solution should turn from a cloudy blue to a clear blue.

NOTE: This step is time-sensitive!! Proceeed to the next step within 2 minutes.

Separate plasmid DNA from denatured proteins and chromosomal DNA

3. Add 350 μL of cold Yellow Zyppy Neutralization buffer (w/RNAase A) to the tube, and mix the contents thoroughly by inverting several times. The solution will turn yellow when neutralization is complete, and a yellowish precipitate will form. Invert the sample an additional 3-4 times to ensure complete neutralization. Be sure there is no more blue color.

4. Centrifuge the mixture at maximum speed for 3 minutes to remove denatured proteins and chromosomal DNA. Notice that the tube contains a white precipitate that has collected on one side of the tube. The pale yellow supernatant contains the plasmid DNA.

Purify plasmid DNA by adsorption to a silica resin.

5. Using a pipette, carefully transfer the pale yellow supernatant (~900 μL) onto a Zyppy spin column. Be careful not to transfer any of the yellow precipitate! Label the column - NOT the collection tube that you will use in the next step.

6. Place the column with the collection tube attached into a centrifuge and spin at maximum speed for ~ 1 minute.

7. Remove the column and discard the flow through in the collection tube.

8. Place the column back into the collection tube and add 200 μL of Zyppy Endo-Wash solution. (Endo-Wash contains guanidine hydrochloride and isopropanol, which will remove denatured proteins from the resin.)

9. Centrifuge for ~1 minute and discard the flow through.

10. Place the column back into the collection tube then add 400 μL of Zyppy Column Wash buffer. (This steps removes contaminating salts.) Centrifuge for ~1 minute. Empty the collecting tube.

11. Repeat the centrifugation step to remove any residual ethanol.

Elute the plasmid DNA

12. Transfer the Zyppy column to a clean (and appropriately labeled) 1.5 mL centrifuge tube, leaving the lid of the tube open.

13. Carefully, add 100 μL of elution buffer directly on top of the white column bed. Place the pipette tip as close as you can to the white column bed without poking it. Slowly dispense the buffer on top of the resin bed.

14. Allow the buffer to percolate into the column by letting the column sit in the microcentrifuge tube for 1 minute.

15. Centrifuge the column at maximum speed for 30 seconds. Again, it’s fine to leave the cap open during this spin.

16. Remove the column, cap the tube and place it on ice. This tube should now contain plasmid DNA. Label the tube. SAVE the DNA for future experiments.


Integrating internet assignments into a biochemistry/molecular biology laboratory course

A main challenge in educating undergraduate students is to introduce them to the Internet and to teach them how to effectively use it in research. To this end, an Internet assignment was developed that introduces students to websites related to biomedical research at the beginning of a biochemistry/molecular biology laboratory course. The basic sites introduced cover many subjects, include searching for specific DNA sequences, restriction enzyme mapping, RNA folding, analyzing three-dimensional protein images, and literature searches. These newly learned Internet skills are incorporated into other aspects of the laboratory course. In the example illustrated here, students sequence DNA inserts from randomly chosen bacterial colonies prepared from a human cDNA plasmid library. The obtained DNA sequence is used to search on-line data bases introduced in the initial Internet assignment to determine whether the cDNAs have already been identified. The sequencing “wet-lab” module, coupled with the Internet assignment, gives students experience in four areas. 1) purifying plasmid DNA, 2) performing restriction endonuclease digests on the plasmid DNA, 3) sequencing double-stranded DNA, and 4) comparison of obtained sequence data to known DNA sequences in the human genome data base in a way that simulates a research experience.

Preparing undergraduate biochemistry and molecular biology students for productive careers in the 21st century requires much more than traditional classroom instruction. Students need hands-on experience in asking relevant research questions and formulating viable hypotheses. Students must also learn how to find, understand, and critically evaluate research already performed, as well as be able to access data bases of information. In the past few years this information has become widely available through the Internet. Students need to be taught how to effectively and efficiently use the Internet as soon as possible in the educational process.

One of my teaching aims has been to organize a course that would enable students to learn how to work in a research setting and then be able to use these skills to work in a research laboratory on or off campus. To that end, I developed Chem 486 (Nucleic Acid Laboratory see Ref. 1 for additional information). One of the first assignments given is to use the Internet to retrieve various types of information regarding genes and gene products. Various wet labs are performed throughout the semester. In the example given here, a wet-lab teaching module is presented in which students learn the principles of plasmid DNA preparation and manipulation, cDNA library principles, and DNA sequencing. Using the data obtained from DNA sequencing, the students search the human genome data base to determine whether the sequenced cDNA has already been identified. The main point of this exercise is to show students that they can generate sequence information from unknown cDNA clones and use the sequencing information to search data bases accessible through the Internet.


Introduction

Proteins are the basic building blocks of life, involved in almost all biochemical reactions in life and are considered as one of the most versatile macromolecules discovered 1 . Proteins can function as a biological catalyst (e.g. amylase), a transportation medium (e.g. haemoglobin), or even as a structural component of mechanical scaffoldings (e.g. skeletal muscle) of an organism. Proteins contain a broad range of functional groups on their amino acids monomers. Thiols, thioethers, carboxamides, alcohol, and carboxylic acids are all commonly present in protein molecules 2 . The arrangement of the functional groups usually determines the final properties of the protein. One interesting group of proteins contains a large number of the hydrophobic functional group that allows them to form a transmembrane protein motif capable of transferring signals from the extracellular matrix into the cell body (Fig. 1) 3 . This is especially important to cells as it facilitates detection of the presence of a pathogen before any infection occur 4 .

The NBS and LRR detect effector proteins from pathogens. The NBS–LRR protein also often has a TIR or CC domain at its N-terminus important for the activation of the immune response. Infection with pathogens transfers protein-mediated signals into the plant cell which activates the R protein and leads to the activation of effector-triggered immunity. TIR, Toll/interleukin-1 receptor-like domain CC, coiled-coil domain NBS, nucleotide binding site LRR, leucine-rich repeats R protein, resistance protein. Figure was reconstructed from Spoel and Dong 3 .

The structural significance of proteins is included in many science-based educational materials. In the fields of chemistry, biochemistry, and molecular biology, core functions of protein structure are often explained as abstract concepts. Hence, students may often face difficulties in understanding fundamental concepts of protein beyond the commonly used “Locks and Keys” concept 5 . Deeper understanding and a better appreciation of the concepts involved in many protein–protein interactions can be achieved by providing a hands-on experience 6 and relating it to actual research applications.

An important, relevant real-world application involves the detection and identification of conserved protein motifs that correlate with the expression of disease resistance genes in crop plants. There are several motif structures that are responsible for the signal transduction system of plant immune systems 7, 8 . This paper focuses on two conserved protein motifs, the characteristic nucleotide binding site (NBS) motif and the leucine-rich repeats (LRR) motif. The NBS and LRR motifs are connected as a complex that mediates the effector-triggered immunity in plants (Fig. 1). The NBS motif is responsible for the activation of kinases in signal transduction pathways 9 while the LRR domain functions in ligand binding and pathogen recognition of the resistance proteins 10 .

Overall sequences among members of the (NBS–LRR) resistance gene family are subjected to many variations and could not be detected directly by cross-hybridization technique. However, short stretches of protein sequences within the gene are well conserved between most resistance genes 8, 9 . These conserved motifs enable the use of polymerase chain reaction (PCR) based technique for the isolation of resistance gene analogues (RGA) from known resistant plants or for use as markers to screen breeding material for potentially resistance/susceptible plants. Utilizing degenerate primers, which are a mixture of primers with different substitutions at selected nucleotides targeted at the NBS domain, is possible to amplify new NBS–LRR genes based on the sequence homology. Here, we report a comprehensive laboratory exercise encompassing both molecular techniques and bioinformatics programs for protein sequence analysis that includes total RNA isolation, PCR amplification of targeted cDNA sequences corresponding to the protein motifs of plant disease resistance genes, and agarose gel visualization of PCR amplicons from the wild relative of black pepper, Piper colubrinum.


AP Lab 3 Sample 3 Mitosis

Lab 3 Mitosis and Meiosis

All new cells come from previously existing cells. New cells are formed by karyokinesis- the process in cell division which involves replication of the cell’s nucleus and cytokinesis-the process in cell division which involves division of the cytoplasm. Two types of nuclear division include mitosis and meiosis. Mitosis typically results in new somatic, or body, cells. Mitotic cell division is involved in the formation of an adult organism from a fertilized egg, asexual reproduction, regeneration, and maintenance or repair of body parts. Meiosis results in the formation of either gametes in animals or spores in plants. The cells formed have half the chromosome number of the parent cell.

Mitosis is best observed in cells that are growing at a rapid pace, such as in the whitefish blastula or onion root cell tips. The root tips contain a special growth region called the apical meristem where the highest percentage of cells are undergoing mitosis. The whitefish blastula is formed immediately after the egg is fertilized, a period of rapid growth and numerous cell divisions where mitosis can be observed.

There are several stages included in before, during, and following mitosis. Interphase occurs right before a cell enters mitosis. During interphase, the cell will have a distinct nucleus with one or more nucleoli, which is filled with a fine network of threads of chromatin. During interphase, DNA replication occurs. After duplication the cell is ready to begin mitosis. Prophase is when the chromatin thickens until condensed into distinct chromosomes. The nuclear envelope dissolves and chromosomes are in the cytoplasm. The first signs of the microtubule-containing spindle also begin to appear. Next the cell begins metaphase. During this phase, the centromere of each chromosome attaches to the spindle and are moved to the center of the cell. This level position is called the metaphase plate. The chromatids separate and pull to opposite poles during the start of anaphase. Once the two chromatids are separate, each is called a chromosome. The last stage of mitosis is telophase. At this time, a new nuclear envelope is formed and the chromosomes gradually uncoil, forming the fine chromatin network seen in interphase. Cytokinesis may occur forming a cleavage furrow that will form two daughter cells when separated.

Meiosis is more complex than mitotic stages and involves two nuclear divisions called Meiosis I and Meiosis II. They result in the production of four haploid gametes and allow genetic variation because of crossing over of genetic material. Prior the process, interphase replicates the DNA. During prophase I, the first meiotic stage, homologous chromosomes move together to form a tetrad and synapsis also begins. This is where crossing over occurs, resulting in the recombination of genes. In Metaphase I, the tetrads move to the metaphase plate in the middle of the cell as on mitotic metaphase. Anaphase I brings the tetrads back to their original two stranded form and moves them to opposite poles. During Telophase I, the centriole is finished and the cell prepares for a second division. In Meiosis II, in Prophase II, centrioles move to opposite ends of the chromosome group. In Metaphase II, the chromosomes are centered within the center of each daughter cell. Anaphase II involves the centromere of the chromatids separating. Telophase II occurs when the divided chromosomes separate into different cells, known as haploid cells.

Sordaria fimicola, an ascomycete fungus, can be used to demonstrate the results of crossing over during meiosis. It spends most of its life haploid and only becomes diploid when the fusion of the mycelia of two different strains results in the fusion of two different types of haploid nuclei to form a diploid nucleus. Meiosis, followed by mitosis, in Sordaria results in the formation of eight haploid ascospores contained within a sac called an ascus. They are contained in a perithecium, a fruiting body, until mature enough to be released. The arrangement of spores directly reflects whether or not crossing over occurred. If an ascus has four tan ascospores in a row and four black ascospores in a row -4:4 arrangement, then no crossing over has taken place. If the asci has black and tan ascospores in sets of two -2:2:2:2 arrangement, or two pairs of black ascospores and four tan ascospores in the middle -2:4:2 arrangement, then crossing over has taken place.

The stages of mitosis can be examined in whitefish blastula and onion root cell tips by using a microscope. The process of crossing over and the stages of meiosis only occur during the creation of gametes and spores.

The materials necessary for this exercise are a light microscope, prepared slides of whitefish blastula, onion root cell tips, pencil, and paper.

For this portion of the lab, materials needed are a bag of color-coded connecting beads and magnetized “centromeres,” several trays, and labels marked interphase, prophase, metaphase, anaphase, and telophase.

Exercise 3A.1: Observing Mitosis

During this experiment, prepared slides of whitefish blastula and onion root tips should be observed under the 10X and 40X objectives of a light microscope. A cell in each stage of mitosis should be identified and sketched.

Exercise 3A.2: Time for Cell Replication

In this section of the lab, use the highest power objective on the microscope to observe and count every cell in the field of view. The cells should be counted according to the stage of mitosis they are in. At least 200 cells and 2 fields of view should be examined and counted. The percentage of cells in each stage is then recorded and the amount of time spent in each phase is calculated.

Exercise 3B.1: Simulation of Meiosis

For this portion of the experiment, a chromosome simulation kit will be used to demonstrate meiosis. Two sets of two strands with each set a different color, are connected to simulate DNA replication in both of the homologous pairs, the stage called interphase. Next, the chromosomes were entwined to represent synapsis in the stage known as prophase. Sections of beads were entwined between the pairs as in crossing over and aligned at the equator. Beads of each pair exchange places, representing metaphase. Next, anaphase was simulated by the homologous pairs being separated to opposite sides of the tray, or in terms of the “chromosomes,” the cell. Pushing the chromosomes into two separate cells, or trays, mimicked telophase.

Meiosis II was simulated as well. Prophase II is shown by the separation of the two beads, but no true change. The chromosomes again move to the equator during metaphase II, and in anaphase II, the two chromatids are separated and moved to opposite poles. Telophase II separates the chromosomes into four different cells.

Exercise 3B.2: Crossing Over during Meiosis in Sordaria

Prepared slides of Sordaria fimicola were observed under a light microscope. The asci were identified as either 4:4 or asci showing crossover. These readings were recorded. The percentage of each and map units were calculated.

Why is it more accurate to call mitosis “nuclear replication” rather than “cellular division”? It is more accurate to describe mitosis as “nuclear replication” because the cell does not divide in any of the mitotic steps. The entire process of mitosis is a series of steps that divides the nucleus into two separate nuclei at opposite poles. When a cell is truly split, the process is known as cytokinesis.

Explain why the whitefish blastula and onion root tips are selected for a study of mitosis. The blastula is what is formed directly following fertilization and, therefore, the cell is growing and many of the phases can be seen at this time. Onion root tip cells are also specimens that include a large amount of cell growth and a high percentage of cells experiencing mitotic activities.

Table 1: Number of Cells in Each Stage of Mitosis and Amount of Time Spent in Each Stage


Results

Pbk was identified as a key protein for mediating HFD-induced compensatory beta cell proliferation using a Pbk kinase dysfunctional mouse model. Mechanistically, the transcription factor JunD recruits menin and HDAC3 complex to the Pbk promoter to reduce histone H3 acetylation, leading to epigenetic repression of Pbk expression. Pharmacologically blocking the menin–JunD interaction menin inhibitors (MIs) increased compensatory beta cell proliferation, resulting in both improved hyperglycemia and glucose tolerance in HFD-induced diabetic mice, demonstrating the key impact of MIs on influencing expression of Pbk and beta cell proliferation in mouse models.


Background

Electrophoresis

As you know from the tyrosinase PAGE lab, electrophoresis is the process of separating particles in an electric field. Like PAGE, nearly all of the common forms of electrophoresis conduct separation in a semisolid gel matrix in which the gel consists of an aqueous phase (buffer) and a solid phase composed of a natural or artificial polymer (agarose, polyacrylamide, starch, etc.). The solid phase has two roles: (1) it acts as a "sieve" to separate molecules according to a specific physical or chemical property and (2) it acts as a "trap" to keep separated molecules from diffusing at the conclusion of the electrophoretic treatment.

In an electric field E, a particle of charge q experiences a force (F) of F = qE. That is, a negatively charged molecule will "feel" a force pushing it against the positive-to-negative electric field lines. Any negatively charged molecule migrates toward the "anode" (+) end of the field, while positively charged molecules migrate toward the "cathode" (-) end.

Since DNA is negatively charged, it migrates towards the anode. The rate at which a particular DNA fragment migrates through a gel is determined by its size. The larger the molecule, the more interaction it has with the gel matrix and the slower it moves. Smaller objects slip through the pores of the gel easily, while larger ones become trapped and move through more slowly. The distance traveled during a defined length of time is inversely proportional to the size of the fragment. The actual size of the pores in the gel influences the speed of migration, and thus this can be adjusted by varying the concentration of the matrix in the gel. Agarose, because it is safe, effective, and easy to handle, is the matrix of choice for gels for basic DNA analysis.

Visualization of DNA

After electrophoresis is completed, the DNA must be stained in order to visualize the separated restriction fragments. The most common method of direct staining is the incubation of the gel in a weak solution of ethidium bromide (EtBr). This compound has UV absorbance maxima at 300 and 360 nm, and can also absorb energy from nucleotides excited at 260nm. This absorbed energy is re-emitted as orange/yellow fluorescence at 590nm. The closer the UV illumination is to 260 nm, the more likely you are to damage the DNA. Illumination

300 nm gives the strongest fluorescence, however, many researchers illuminate with 360 nm to protect their sample from damage (360 nm gives considerably weaker fluorescence). Ethidium cations have high affinity for binding DNA. In their presence, the DNA dynamically opens a space between its base pairs by unwinding. The ethidium cation unwinds DNA by about 26°. The ion fits into a 0.34 nm (3.4 Å) opening created by this unwinding that is stabilized by hydrophobic interactions between ethidium and the nucleotide bases (see picture). This unwinding induces local structural changes to the DNA strand, such as lengthening of the DNA strand, or twisting of the base pairs. These structural modifications can lead to functional changes, often to the inhibition of transcription and replication and DNA repair processes, which makes intercalators potent mutagens. For this reason, DNA intercalators are generally considered carcinogenic and should be handled with care.

Restriction mapping of plasmid DNA

A plasmid is a DNA molecule that is separate from, and can replicate independently of, the chromosomal DNA. They are double stranded and, in most cases, circular. Plasmids usually occur naturally in bacteria, but are sometimes found in eukaryotic organisms. Last week, we set-up a hypothetical scenario in which the labels have rubbed off of tubes of E. coli each carrying a plasmid that your lab uses for research. The four possible plasmid restriction maps are given below (images are also available on the D2L main page).

On each map, you see the relative locations for where various restriction enzymes cut. The arrows indicate the locations of important genetic features. "AmpR" expresses a protein called beta-lactamase, which confers ampicillin resistance to the bacterium carrying the plasmid. The ampicillin resistance trait is used to select plasmid-containing bacteria that grow in the presence of this antibiotic. The region labeled "ori" is the E. coli "origin of replication," a sequence to which DNA polymerase binds. The origin of replication is required for E. coli to pass the plasmid to daughter cells. The "rop" gene expresses a small protein that helps maintain a high copy number of about 20 plasmids per cell. This feature ensures that we are able to isolate more DNA per cell. The other features express proteins that are of interest to your lab in this hypothetical scenario. They were inserted into the plasmids using methods described in Lehninger Section 9.1.

Look carefully at the differences between the four plasmids. We will use the pattern that results from reaction with restriction enzymes to identify our plasmid. As an example, of what we may see, digests by either EcoRI or AvaI will simply give you a piece that is the size of the entire plasmid because there is only one cut-site for each of them. However, a double digest with both EcoRI and AvaI will allow you to distinguish pAB125 from the others. Plasmid pAB125 will give a large fragment

4000 bp in length and a small fragment

300 bp in length. In contrast, the smaller fragment resulting from a double digest with the other three plasmids would be a clearly distinguishable

Last week you performed a small-scale isolation of plasmid DNA. This week you will map the restriction sites of four available enzymes: EcoRI, AvaI, HincII, and RsaI. You will use the lengths of the DNA fragments resulting from your restriction digests to map their relative locations of the sites on the plasmid. With this information, you will identify your "unknown" plasmid from the four possibilities. You will also complete the map by assigning position numbers to the sites.

Your agarose gel will have a total of ten lanes. Be sure to run appropriate controls on each gel, including single digests to compare with double digests or double digests to be compared with each other. The more digests you have on your gel, the more information you will obtain by comparing fragment sizes directly. You will generate a standard curve of fragment sizes by running a DNA ladder mixture on each gel. Our reference DNA mixture will be "2-log DNA Ladder" (New England Biolabs product N3200). Visit the manufacturer's website for more information on the composition of this DNA reference mixture (link on the D2L main page).

Restriction Endonuclease Enzymes

So what is a restriction enzyme? These enzymes are endonucleases (polynucleotide degrading enzymes) that recognize a specific DNA sequence, and introduce double-stranded cuts. Researchers stumbled upon these enzymes while observing the growth of a bacteriophage virus, and saw that its growth was "restricted" in some bacterial cultures. Upon investigation they found that in these cultures, endonucleases degraded the unmethylated viral DNA upon entry into the bacterial cell, restricting phage growth (hence the name restriction enzyme). 1 Methylation of the bacteria's own DNA prevents the enzymes from destroying host DNA, while allowing them to target any foreign DNA that invades the cell.

As can be seen in the table above, there are a multitude of restriction enzymes, each recognizing its own "cut-sequence." These recognition sites are typically 4-8 bases in length, and are usually palindromic (the 5'-3' sequence is the same for top and bottom strands). In the above table, the arrows indicate where the restriction enzyme cleaves the phosphodiester bond in the DNA backbone, with two bonds being hydrolyzed in order to achieve the double-stranded cut. The resulting, cut DNA strands can either form blunt ends (no unpaired bases) or sticky ends (multiple unpaired bases). Sticky ends are of particular use due to their abilities to hydrogen-bond with complementary sequences digested with the same enzyme. As we will see later, this plays a large role in restriction based DNA cloning.

The enzyme shown on the right, EcoRI, gives insight into the palindromic nature of the sequence recognition sites. Restriction enzymes are homodimers, with each subunit recognizing the same sequence of DNA on each strand. The purple balls in this model are Mg 2+ ions, which, as you can recall from last week, are essential in stabilizing the negative charges in the enzyme-DNA reaction complex.

Cloning with Restriction Enzymes

It is no understatement to say that the discovery of restriction enzymes revolutionized biochemistry and molecular biology. Advances in DNA technology quickly followed the discovery of restriction enzymes: cloning, DNA fragmentation analysis, and DNA restriciton mapping are only some of the many applications of restriction enzymes. 2

Cloning involves the combination of two or more sets of DNA into one, recombinant piece of DNA. In terms of creating recombinant plasmids, the two DNA pieces are usually a vector backbone (1) and a gene insert (2). The insert encodes for the protein, enzyme, or function of interest while the backbone allows for transformation of the plasmid into bacterial cells, and for replication of the plasmid upon transformation.

In order to perform this type of cloning, both the vector and the insert must be digested with the same enzyme(3). This produces sticky ends on both the vector and the insert which will overlap with each other when combined. The pieces are loosely held in place by these sticky ends, and are ligated together with DNA ligase, which re-forms the phosphodiester bonds in the DNA backbone. The ligated plasmid can then be transformed into a host cell (4), and will copy itself using the replication machinery of the host (5).

This method may seem complicated, but it can be thought of in fairly simple terms. Restriction enzymes act as molecular scissors which cut DNA sequences from two or more sources. The sticky ends generated from these cuts act as a weak adhesive that reversibly associate the insert to the vector. Finally, the DNA ligase acts as a laminate, sealing the two pieces of DNA together with a covalent bond that can not be easily reversed.

DNA Fragmentation Analysis and Plasmid Mapping

A second application for restriction enzymes in biochemistry is their ability to "map" segments of DNA through their patterns of cleavage. Information about the size and relative locations of restriction cut sites can be found by digesting DNA with a series of restriction enzymes. Running the products of a DNA restriction digest on an electrophoresis gel will allow visualization of the cuts, as the single, large DNA band will turn into multiple, smaller digested bands. This mapping is what you will be carrying out on the unknown plasmid in Lab 8.

How to approach plasmid mapping will be covered in more detail in the Data Analysis section.

From the gel image, the lengths of the bands can be determined by measuring the distance each band has travelled. First, the migration distances of the ladder bands should be measured and plotted in Excel against the 10 of the number of base pairs. The image to the left gives an indication of how to measure the distances. Make sure you are consistent with your measurements (i.e. start at the bottom of each well, and end at the middle of each band).


Transduction and Analysis

Once you have generated a lentiviral stock with a suitable titer, you are ready to transduce the lentiviral construct into the mammalian cell line of choice and assay for expression of your recombinant protein. Guidelines are provided below.

Your lentiviral construct contains a deletion in the 3′ LTR that leads to self-inactivation of the lentivirus after transduction into mammalian cells. Once integrated into the genome, the lentivirus can no longer produce packageable virus.

Transient vs. Stable Expression

After transducing your lentiviral construct into the mammalian cell line of choice, you may assay for expression of your gene of interest in the following ways:

  • Pool a heterogeneous population of cells and test for expression directly after transduction (i.e. “transient” expression). Note that you must wait for a minimum of 48–72 hours after transduction before harvesting your cells to allow expressed protein to accumulate in transduced cells.
  • Select for stably transduced cells using Blasticidin or Zeocin, as appropriate. This requires a minimum of 10-12 days after transduction, but allows generation of clonal cell lines that stably express the gene of interest. Note: We have observed stable expression of a target gene for at least 6 weeks following transduction and selection.

Determining Antibiotic Sensitivity for Your Cell Line

If you wish to select for stably transduced cells, you must first determine the minimum concentration of Blasticidin or Zeocin, as appropriate, required to kill your untransduced mammalian cell line (i.e. perform a kill curve experiment). If you titered your lentiviral construct in the same mammalian cell line that you are using to perform your stable expression experiment, then you may use the same concentration of Blasticidin or Zeocin for selection that you used for titering.

Multiplicity of Infection (MOI)

To obtain optimal expression of your gene of interest, you will need to transduce the lentiviral construct into your mammalian cell line of choice using a suitable MOI. MOI is defined as the number of virus particles per cell and generally correlates with the number of integration events and as a result, expression of your gene of interest. Typically, expression levels increase linearly as the MOI increases.

Determining the Optimal MOI

A number of factors can influence optimal MOI including the nature of your mammalian cell line (e.g. non-dividing vs. dividing cell type see Recommendation below), its transduction efficiency, your application of interest, and the nature of your gene of interest. If you are transducing your lentiviral construct into the mammalian cell line of choice for the first time, we recommend using a range of MOI (e.g. 0, 0.5, 1, 2, 5, 10) to determine the MOI required to obtain the optimal expression of your protein for your application.

In general, we have found that 80-90% of the cells in an actively dividing cell line (e.g. HT1080) express a target gene when transduced at an MOI of

1. Some nondividing cell types transduce lentiviral constructs less efficiently. For example, only about 50% of the cells in a culture of primary human fibroblasts express a target gene when transduced at an MOI of

1. If you are transducing your lentiviral construct into a non-dividing cell type, you may need to increase the MOI (e.g. MOI = 10) to achieve optimal expression levels for your recombinant protein.

Control lentiviral vectors expressing lacZ or EmGFP are available for optimization. If you have generated a lentiviral stock of a lacZ expression control (e.g. pLenti6/V5-GW/lacZ), we recommend using the stock to help you determine the optimal MOI for your particular cell line and application. Once you have transduced the control lentivirus into your mammalian cell line of choice, the gene encoding β-galactosidase or EmGFP will be constitutively expressed and can be easily assayed (refer to the expression vector or expression control vector manual for assay methods).

Viral supernatants are generated by harvesting spent media containing virus from the 293FT producer cells. Spent media lacks nutrients and may contain some toxic metabolic waste products. If you are using a large volume of viral supernatant to transduce your mammalian cell line (e.g. 1 ml of viral supernatant per well in a 6-well plate), note that growth characteristics or morphology of the cells may be affected during transduction. These effects are generally alleviated after transduction when the media is replaced with fresh, complete media.

You will need the following items:

  • Your titered lentiviral stock (store at –80°C until use)
  • Mammalian cell line of choice
  • Complete culture medium for your cell line
  • 6 mg/ml Polybrene, if desired
  • Appropriately sized tissue culture plates for your application
  • Blasticidin or Zeocin, as appropriate (if selecting for stably transduced cells)

Transduction Procedure

Follow the procedure below to transduce the mammalian cell line of choice with your lentiviral construct. Reminder: If you are performing Zeocin selection, remember that cells should not be confluent at the time of selection (see Step 6 below). Plate your cells accordingly.

  1. Plate cells in complete media as appropriate for your application.
  2. On the day of transduction (Day 1), thaw your lentiviral stock, and if necessary, dilute the appropriate amount of virus into fresh complete medium to obtain a suitable MOI. Keep the total volume of medium containing virus as low as possible to maximize transduction efficiency. DO NOT vortex.
  3. Remove the culture medium from the cells. Mix the medium containing virus gently by pipetting and add to the cells.
  4. Add Polybrene® (if desired) to a final concentration up to 10 μg/ml. Swirl the plate gently to mix. Incubate at 37° in a humidified 5% CO2 incubator overnight. Note: If you are transducing cells with undiluted viral stock and are concerned about possible toxicity or growth effects caused by overnight incubation, it is possible to incubate cells for as little as 6 hours prior to changing medium.
  5. The following day (Day 2), remove the medium containing virus and replace with fresh, complete culture medium. Incubate at 37°C in a humidified 5% CO2 incubator overnight.
  6. The following day (Day 3), perform one of the following:
    • Harvest the cells and assay for expression of your recombinant protein if you are performing transient expression experiments.
    • Remove the medium and replace with fresh, complete medium containing the appropriate amount of Blasticidin or Zeocin™, as appropriate to select for stably transduced cells. Proceed to Step 7.

Integration of the lentivirus into the genome is random. Depending upon the influence of the surrounding genomic sequences at the integration site, you may see varying levels of recombinant protein expression from different antibiotic-resistant clones. We recommend testing at least 5 antibiotic-resistant clones and selecting the clone that provides the optimal expression of your recombinant protein for further studies.

Detecting Recombinant Protein

You may use any method of choice to detect your recombinant protein of interest including functional analysis, immunofluorescence, or western blot. If you have cloned your gene of interest in frame with an epitope tag, you may easily detect your recombinant protein in a western blot using an antibody to the epitope tag


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Extending the small-molecule similarity principle to all levels of biology with the Chemical Checker

Small molecules are usually compared by their chemical structure, but there is no unified analytic framework for representing and comparing their biological activity. We present the Chemical Checker (CC), which provides processed, harmonized and integrated bioactivity data on

800,000 small molecules. The CC divides data into five levels of increasing complexity, from the chemical properties of compounds to their clinical outcomes. In between, it includes targets, off-targets, networks and cell-level information, such as omics data, growth inhibition and morphology. Bioactivity data are expressed in a vector format, extending the concept of chemical similarity to similarity between bioactivity signatures. We show how CC signatures can aid drug discovery tasks, including target identification and library characterization. We also demonstrate the discovery of compounds that reverse and mimic biological signatures of disease models and genetic perturbations in cases that could not be addressed using chemical information alone. Overall, the CC signatures facilitate the conversion of bioactivity data to a format that is readily amenable to machine learning methods.


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